Dynamics of the Lipid Droplet Proteome of the Oleaginous YeastRhodosporidium toruloides

Zhiwei Zhu,a Yunfeng Ding,b Zhiwei Gong,a Li Yang,b Sufang Zhang,a,c Congyan Zhang,b Xinping Lin,a Hongwei Shen,a,c Hanfa Zou,a

Zhensheng Xie,b Fuquan Yang,b Xudong Zhao,b Pingsheng Liu,b Zongbao K. Zhaoa,c

Division of Biotechnology, Dalian Institute of Chemical Physics, CAS, Dalian, Chinaa; National Laboratory of Biomacromolecules, Institute of Biophysics, CAS, Beijing,Chinab; Dalian National Laboratory for Clean Energy, Dalian Institute of Chemical Physics, CAS, Dalian, Chinac

Lipid droplets (LDs) are ubiquitous organelles that serve as a neutral lipid reservoir and a hub for lipid metabolism. Manipulat-ing LD formation, evolution, and mobilization in oleaginous species may lead to the production of fatty acid-derived biofuelsand chemicals. However, key factors regulating LD dynamics remain poorly characterized. Here we purified the LDs and identi-fied LD-associated proteins from cells of the lipid-producing yeast Rhodosporidium toruloides cultured under nutrient-rich,nitrogen-limited, and phosphorus-limited conditions. The LD proteome consisted of 226 proteins, many of which are involvedin lipid metabolism and LD formation and evolution. Further analysis of our previous comparative transcriptome and proteomedata sets indicated that the transcription level of 85 genes and protein abundance of 77 proteins changed under nutrient-limitedconditions. Such changes were highly relevant to lipid accumulation and partially confirmed by reverse transcription-quantita-tive PCR. We demonstrated that the major LD structure protein Ldp1 is an LD marker protein being upregulated in lipid-richcells. When overexpressed in Saccharomyces cerevisiae, Ldp1 localized on the LD surface and facilitated giant LD formation, sug-gesting that Ldp1 plays an important role in controlling LD dynamics. Our results significantly advance the understanding of themolecular basis of lipid overproduction and storage in oleaginous yeasts and will be valuable for the development of superiorlipid producers.

Lipid droplets (LDs), intracellular organelles with deposits ofneutral lipids and involved in many cellular activities, are

widely present in both eukaryotic and prokaryotic cells (1–4).These organelles consist of a neutral lipid core surrounded by aphospholipid monolayer and associated proteins (3, 5). It hasbeen known that LDs serve as the energy reservoir of cells, whichmay increase the adaptation by mobilization and degradation oflipids during nutrient deprivation, and also connect with othercellular processes, including lipid transport, membrane biogene-sis, lipotoxicity relief, protein storage and degradation, pathoge-nicity, and autophagy (6–9). Because the biology of LDs is closelylinked to some diseases, such as obesity, type 2 diabetes, andatherosclerosis, great progress has been made in elucidating thecellular trafficking, dynamics, and biogenesis of LDs in mamma-lian cells (2, 7, 10). However, there have been few studies on LDs inother species, especially naturally lipid-producing microorgan-isms (11–13). Analysis of these microorganisms is motivated bythe fact that microbial lipid production holds a great promise toconvert waste materials, including lignocellulosic biomass, intofatty acid-derived fuel molecules and chemicals in a scenario ofbiorefinery and sustainable development (14, 15).

The major components of LDs are neutral lipids, includingtriacylglycerols (TAGs), sterol esters, and ether lipids (16). Neu-tral lipids constitute more than 90% of LDs by weight, but theratio of TAGs to sterol esters can change substantially dependingon culture conditions and organisms (13, 17, 18). Polar lipids,mainly phosphatidylcholine, phosphatidylethanolamine, andphosphatidylinositol, form a micellar structure that stabilizes thehydrophobic center. Besides lipids, purified LDs have also beenfound to be associated with hundreds of proteins (10). The PATfamily proteins are present on the LD surface in evolutionarilydistant organisms (19). Other LD proteins include lipid metabo-lism enzymes, Rab GTPases, coatomer components, ARF-related

proteins, and SNARE proteins (20). The presence of functionallydiversified proteins in LDs suggests that the biological function ofLDs is much more than just an energy reservoir (10, 21).

The basidiomycetous yeast Rhodosporidium toruloides is robustin terms of lipid production under nitrogen-limited (NL) or phos-phorus-limited (PL) conditions (15, 22, 23). When R. toruloidescells accumulated substantial lipids, giant LDs occupied almostthe entire cells. Our previous analysis of soluble cell lysates of R.toruloides showed that the levels of over 500 proteins were signif-icantly changed during lipid accumulation in response to nitrogenlimitation (24). However, the identity and dynamics of LD-asso-ciated proteome remained unknown in this and other basidiomy-cetous yeasts.

In this study, we cultured R. toruloides in nutrient-rich, NL,and PL media, obtained cell samples of different lipid contents,purified LDs, and identified 226 LD-associated proteins. Theseproteins had diverse biological functions and were involved inlipid metabolism, vesicle traffic, small molecule metabolism, pro-

Received 16 June 2014 Accepted 4 January 2015

Accepted manuscript posted online 9 January 2015

Citation Zhu Z, Ding Y, Gong Z, Yang L, Zhang S, Zhang C, Lin X, Shen H, Zou H,Xie Z, Yang F, Zhao X, Liu P, Zhao ZK. 2015. Dynamics of the lipid dropletproteome of the oleaginous yeast Rhodosporidium toruloides. Eukaryot Cell14:252–264. doi:10.1128/EC.00141-14.

Address correspondence to Zongbao K. Zhao, zhaozb@dicp.ac.cn, or PingshengLiu, pliu@ibp.ac.cn.

Z.Z. and Y.D. contributed equally to this article.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/EC.00141-14.

Copyright © 2015, American Society for Microbiology. All Rights Reserved.

doi:10.1128/EC.00141-14

252 ec.asm.org March 2015 Volume 14 Number 3Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

tein synthesis and processing, or mitochondrial function. Furtherreanalysis of our previous differential transcriptome and pro-teome data sets (24, 25) indicated that the expression levels of themajority of LD-associated proteins changed in response to cultureconditions. Such changes were partially confirmed by reversetranscription-quantitative PCR (RT-qPCR) analysis. Interest-ingly, we found that RHTO_05627 was an abundant LD-associ-ated protein, and thus it was termed Ldp1 (lipid droplet protein1). RT-qPCR and quantitative “omic” data indicated that the ex-pression level of LDP1 was higher in cells with more lipids than incells with fewer lipids. Western blot analysis showed that Ldp1 waspresent specifically in the LD fraction, and its abundance in LDsincreased in lipid-rich cells. We showed that overexpressed Ldp1localized on the LD surface and facilitated the formation of giantLDs in engineered ascomycetous yeast Saccharomyces cerevisiaestrains.

This multidisciplinary study advanced the understanding ofthe molecular basis of neutral lipid overproduction and storage inoleaginous yeasts. Such information will facilitate LD research ingeneral as well as the development of superior lipid producers forthe production of biofuels and chemicals.

MATERIALS AND METHODSStrain and culture conditions. R. toruloides CGMCC 2.1389 was obtainedfrom the China General Microbiological Culture Collection (Beijing,China). It was the parent strain of the sequenced haploid NP11 (24). Thestrain in glycerol stock was revived at 28°C on yeast extract-peptone-dextrose (YPD) agar slants (10 g/liter yeast extract, 10 g/liter peptone, 20g/liter dextrose, and 15 g/liter agar, pH 6.0). Inocula were grown in YPDliquid medium at 30°C and 200 rpm for 24 h. Cells were separately cul-tured in nutrient-rich (YPD), NL, or PL medium with 10% (vol/vol)initial inoculum at 30°C and 200 rpm. The NL medium consisted of 70g/liter glucose, 0.1 g/liter (NH4)2SO4, 0.75 g/liter yeast extract, 1.0 g/literKH2PO4, 1.5 g/liter MgSO4 · 7H2O, and 1% (vol/vol) trace element solu-tion, pH 6.0. The PL medium consisted of 70 g/liter glucose, 5 g/liter(NH4)2SO4, 2 g/liter peptone, 2.3 g/liter K2SO4, 1.5 g/liter MgSO4 · 7H2O,and 1% (vol/vol) trace element solution, pH 6.0. The trace element solu-tion contained 4.0 g CaCl2 · 2H2O, 0.55 g FeSO4 · 7H2O, 0.52 g citrate ·H2O, 0.10 g ZnSO4 · 7H2O, 0.076 g MnSO4 · H2O, and 100 �l of 18 MH2SO4 per liter (25). The initial carbon-to-nitrogen (C/N) molar ratios ofYPD, NL, and PL media were 3.9, 309, and 24.8, respectively. The initialcarbon-to-phosphorus (C/P) molar ratios of YPD, NL, and PL mediawere 143, 280, and 10,510, respectively. The culture pH was adjusted to 6.0with 2 M NaOH once per 24 h. Cell pellets harvested from the culturebroth grown in YPD medium at 24 h, in NL medium at 24 h, and in PLmedium at 48 h were subsequently used for LD preparation and RT-qPCRanalysis.

LD purification. LDs were isolated at 4°C according to a previouslydescribed protocol with minor modifications (26). Briefly, cell pellets har-vested from 400 ml of culture broth were washed twice with 30 ml ofphosphate-buffered saline (PBS) buffer and then suspended in 30 ml ofbuffer A (20 mM tricine, 250 mM sucrose, pH 7.8) containing 0.2 mMphenylmethylsulfonyl fluoride (PMSF) and incubated on ice for 20 min.These cell samples were then homogenized by running them four timesover a JN3000-type ultrahigh-pressure continuous flow cell disrupter(JNBIO Co., Ltd., Guangzhou, China) at 150 MPa. For cells cultured inYPD, the cell debris of the homogenate was removed by centrifugation at3,000 � g for 10 min. Eight milliliters of supernatant layered with 2 ml ofbuffer B (20 mM HEPES, 100 mM KCl, 2 mM MgCl2, pH 7.4) on top wascentrifuged in an SW40 rotor at 8,000 � g for 40 min. The floating LDswere collected in a 1.5-ml Eppendorf tube and washed three times withbuffer B. For the cells cultured in NL medium or PL medium, the homog-

enate was centrifuged directly at 4,000 � g for 15 min. The LDs werecollected in 1.5-ml Eppendorf tubes and washed three times with buffer A.

Lipid extraction and analysis. Wet cells were harvested from 30 mlculture broth by centrifugation and washed twice with distilled water. Thecell mass, expressed as cell dry weight (CDW), was determined gravimet-rically after drying the wet cells at 105°C for 24 h. Dried cells were digestedwith 4 M HCl at 78°C for 1 h before extraction with chloroform-methanol(1:1, vol/vol). The extracts were washed with 0.1% NaCl solution, driedover anhydrous Na2SO4, and evaporated in vacuo, and finally the residueswere dried at 105°C for 24 h to yield the total lipids (27). Lipid content wasexpressed as gram lipids per gram CDW. To purify lipids from LDs, sam-ples were extracted by chloroform-acetone (1:1, vol/vol) and dried withhigh-purity nitrogen gas. Neutral lipids and polar phospholipids wereseparated by silica gel thin-layer chromatography with hexane-diethylether-acetic acid (80:20:1, vol/vol/vol) and chloroform-methanol-aceticacid-H2O (75:13:9:3, vol/vol/vol/vol) solvent systems, respectively, andvisualized by iodine vapor (11).

LD protein separation and identification. One milliliter of chloro-form-acetone (1:1, vol/vol) was added to the purified LDs to extract lipidsand precipitate LD proteins. The pellets were harvested by centrifugationat 20,000 � g for 10 min, dissolved in 2� SDS-PAGE sample buffer, andthen denatured at 95°C for 5 min. The total membrane, cytosol, andpostnuclear supernatant (PNS) fractions were obtained by following aprotocol previously described (26). The protein samples of the LD, mem-brane, cytosol, and PNS were separated on a 10% SDS-polyacrylamide geland stained with colloidal blue. For mass spectrometry analysis, the entirelane corresponding to LD proteins from cells cultured in YPD mediumwas cut into 29 slices. In the NL and PL samples, seven bands were slicedbecause they were obviously different from those in the YPD sample.These gel slices were destained twice with 200 �l acetonitrile–25 mMammonium bicarbonate (2:3, vol/vol), dehydrated with acetonitrile, andsubjected to reduction and alkylation treatment as described before (11).Gel slices were washed sequentially with 25 mM ammonium bicarbonate,25 mM ammonium bicarbonate solution–acetonitrile (1:1, vol/vol), andacetonitrile, dried in a SpeedVac vacuum concentrator (ThermoFisherScientific, Waltham, MA, USA), and incubated in trypsin solution (10ng/�l in 25 mM ammonium bicarbonate) at 4°C for 30 min. The excesstrypsin solution was removed, and 50 �l ammonium bicarbonate (25mM) was added for overnight digestion at 37°C. The digestion reactionwas stopped by adding formic acid to a final concentration of 0.1%.

The peptides extracted from in-gel digestion were analyzed by a nano-liquid chromatography-electrospray ionization-linear trap quadrupoletandem mass spectrometry (LC-ESI-LTQ MS/MS) system as describedbefore (11). The peptide solution loaded onto a C18 trap column by anautosampler was eluted onto a C18 column (100 mm by 100 �m) packedwith Sunchrom packing material (SP-120-3-ODS-A, 3 �m) and then sub-jected to an LTQ linear quadrupole ion trap mass spectrometer (Thermo-Fisher Scientific) equipped with an electrospray ion source. The LTQ massspectrometer was operated in a data-dependent mode with the initial MSscan ranging from 400 to 2,000 Da. The five most abundant ions wereselected automatically for subsequent collision-activated dissociation.The normalized collision energy was 35, and the activation time was 30 ms.The dynamic exclusion parameters were as follows: repeat count, 1; repeatduration, 30.00; exclusion list size, 500; exclusion duration, 90.00; exclusionmass width by mass, �1.50 atomic mass units (amu). The peak list files weregenerated by Extract_msn.exe in a BioWorks package (v3.3.1 SP1) with de-fault parameters. All MS/MS data were searched against the R. toruloidesprotein database containing 8,171 proteins (http://www.bioconversion.dicp.ac.cn/EWEB/DATA/02/RT.pep.corrected.annotated.nameCH.fa.rar). It should be noted that contaminants such as trypsin and keratinswere also included in the database. The search parameters of SEQUEST inthe BioWorks suite were set as follows: enzyme, trypsin as specific pro-tease, and two miscleavages allowed; precursor ion mass tolerance, 2.0 Da;and fragment ion mass tolerance, 1.0 Da. The variable modification wasset to oxidation of methionine (Met � 15.99 Da). The fixed modification

Dynamics of R. toruloides Lipid Droplet Proteome

March 2015 Volume 14 Number 3 ec.asm.org 253Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

was set to carboxyamidomethylation of cysteine (Cys � 57.02 Da). Thesearch results were filtered with Xcorr versus Charge values as follows:Xcorr (� 1) � 1.90, Xcorr (� 2) � 2.50, and Xcorr (� 3) � 3.75). Thepeptide assignment probabilities were automatically generated by the Bio-Works package using the SRF files. Proteins with at least two distinctpeptides were assigned as identified proteins.

Protein function prediction and classification. The S. cerevisiae or-thologs of identified proteins were obtained by protein sequence similar-ity searching against the Saccharomyces Genome Database (SGD; http://www.yeastgenome.org/). An E value below 1E�10 was used as athreshold for homologous candidates. Functional classification of the LDproteins was based on protein annotation and SGD gene description. Theorthologs of previously identified LD proteins in humans (21, 28), fly (21,29), S. cerevisiae (17, 21, 30), Pichia pastoris (18), and Yarrowia lipolytica(13) were assigned by reciprocal BLAST.

Differentially expressed transcripts and proteins. The transcript andprotein levels of the genes coding for LD-associated proteins were re-trieved from the database of our previous digital gene expression profilingand semiquantitative proteome study of R. toruloides (24). The sequencetags of digital gene expression profiling (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc�GSE39023) were cleaned and normalized to tagsper million (TPM) and then mapped to annotated transcripts to calculatethe transcription level. Optimal criteria implemented with SFOER soft-ware (31) were used for peptide assignment filter in the proteome analysisof the soluble cell lysates, which made the false-discovery rate below 1%.Spectral counts (http://www.bioconversion.dicp.ac.cn/EWEB/DATA/03/Peptide_identification.zip) of the identified peptides were collected andnormalized as described previously (32). The same statistical algorithmsfor significance tests as those in previous processing (24) were used. Pvalues of 0.01 and fold changes of �2 were used as the thresholds of thedifferentially expressed transcripts, while P values of 0.05, G values of�1.35, and fold changes of �2 were used for assignment of the changedLD proteins.

RT-qPCR analysis. The cell pellets harvested from 1 ml culture brothof R. toruloides were immediately frozen in liquid nitrogen and stored at�70°C. Total RNA was isolated from about 30- to 50-mg cell samplesusing the FastRNA Pro Red kit and FastPrep instrument (Qbiogen, Inc.,Irvine, CA, USA) according to the manufacturer’s instructions, and thesetting of FastPrep Instrument was 6.0 m/s for 60 s. The RNA concentra-tion and quality were determined by a Nanodrop ND1000 spectropho-tometer (ThermoFisher Scientific), while the RNA integrity was assessedby agarose gel electrophoresis. DNA removal and cDNA synthesis wereperformed with the PrimerScript RT reagent kit with gDNA Eraser(TaKaRa, Dalian, China). Equal amounts of each RNA (750 ng) were usedfor reverse transcription. Quantitative PCR (qPCR) experiments werecarried out using the SYBR Premix Ex Taq GC kit (TaKaRa) in an Ecoreal-time PCR system (Illumina, San Diego, CA, USA). Gene-specificprimers are listed in Table S1 in the supplemental material. Reactionswere performed with appropriate amounts of cDNA templates and 0.2�M each primer in a total volume of 10 �l. Nonspecific amplicons werenot observed by dissociation curves and agarose gel electrophoresis anal-ysis. The amplification efficiencies were calculated from five-point cali-bration curves based on a 6-fold dilution series using a pool of equalvolumes of all cDNA as the template (see Table S1 in the supplementalmaterial). The expression levels were normalized to two reference genes(RHTO_03560, coding for actin, and RHTO_03746, coding for glyceral-dehyde 3-phosphate dehydrogenase) by a normalization factor as de-scribed before (33). Three biological and two technical replicates wereanalyzed to obtain the relative expression quantity.

Western blotting. Anti-Ldp1 polyclonal antibody was raised againsttwo synthesized peptides, NEKQPATDAPLAHEC (corresponding toamino acid residues 5 to 18) and CKKKDEVAETVEEKEGEGEK (corre-sponding to amino acid residues 246 to 264), selected from the N and Ctermini of Ldp1. The mixture of these two peptides was used as antigensand injected into two rabbits. After three injections, the antibody activity

of the rabbit sera was confirmed by Western blotting. Four cellular frac-tions (LD, total membrane, cytosol, and PNS) of R. toruloides were pre-pared according to a previously reported protocol (26). Proteins wereseparated by 12% SDS-PAGE, transferred onto polyvinylidene difluoride(PVDF) membranes, blotted with the anti-Ldp1 polyclonal antibody, anddetected with the enhanced chemiluminescence system.

Cloning and expression of LDP1. The cDNA of Ldp1 (NCBI acces-sion number EMS18697) was amplified from the reverse transcriptionproduct with primers PRLP1-L1 (CAAACAACGAGCACAGCGACAC)and PRLP1-R1 (AAACCGAGAAGAAACCCGAAC). The amplificationproduct was first cloned into a pMD-19T vector (TaKaRa) to give pMD-19T:perilipin. The complete open reading frame of Ldp1 was amplifiedwith the primer pair PG-PER-F (GGAATTCAACATGGCCACCGTCAACGAGAAGC), where underlining indicates the EcoRI restriction site, andPG-PER-R (TGCAAGCTTGGCGTAATCATGGTCATCTGCTTC T C C C C C T C G CCCTCCT). The green fluorescent protein (GFP) gene was amplified frompGFPuv (Clontech Laboratories, Inc., Mountain View, CA, USA) with theprimer pair PG-GFP-F (GAGAAGGAGGGCGAGGGGGAGAAGCAGATGACCATGATTACGCCAAGCT) and PG-GFP-R (AAAAGGAAAAGCGGCCGCTCATTTGTAGAGCTCATCCATGCCATG), where underlin-ing indicates the NotI restriction site. The amplification products were gelpurified and spliced by overlap extension PCR (34). The fusion segmentwas digested with EcoRI and NotI and inserted into pYES2/CT (Invitro-gen Co., Carlsbad, CA, USA) to produce the plasmid pYES2-PG express-ing the C-terminal GFP-tagged Ldp1 fusion protein (Ldp1-GFP) underthe control of the GAL1 promoter. pYES2-PG was digested with HindIIIto remove the LDP1 fragment and self-ligated to generate pYES2-GFP.Both plasmids were transformed by electroporation into S. cerevisiae IN-Vsc1 according to a previous protocol (35). The pYES2-PG plasmid wasalso transformed to strains expressing Erg6-RFP or Sec13-RFP (36).

Confocal fluorescence microscopy. Single colonies from the transfor-mation plates were first cultured at 30°C in the maintenance medium (20g/liter raffinose, 6.7 g/liter yeast nitrogen base without amino acids, andsupplementary amino acids as needed, i.e., 100 mg/liter leucine, 20 mg/liter histidine, 20 mg/liter tryptophan, or 30 mg/liter lysine) to an opticaldensity at 600 nm of 3.0. One milliliter of preculture was used to inoculate10 ml of inducing medium (replacement of raffinose with galactose in themaintenance medium) supplemented with 200 �M oleic acid. After cul-tivation at 30°C for 30 h, cells were collected and washed twice with PBSand then dropped onto microscope slides pretreated with collagen for 20min. The samples were stained with a 500-fold-diluted solution of Lipid-TOX Red (catalog number H34477; Invitrogen) in darkness for 20 minand viewed with an Olympus FV1000 confocal microscope (OlympusCo., Tokyo, Japan).

RESULTSLipid production and LD morphology under different cultureconditions. It has been known that oleaginous yeasts accumulatelipids under nutrient-limited conditions and that cellular lipidcontents are low (below 20%) under nutrient-rich conditions(22). In the present study, the lipid contents of the inoculum cellsand the cells cultured in YPD medium for 24 h were 8.6% and8.2%, respectively; however, lipid content increased to 51.5%when the cells were grown in NL medium for 72 h (NL-72 h).Similarly, in PL medium, the cells accumulated lipids to 22.3%and 31.7% after 48 h and 72 h, respectively (Fig. 1A), indicatingthat the cells grew and accumulated lipids slower in PL medium.The lipid accumulation was consistent with the evolution of LDsize. Several small LDs were present in low-lipid cells, while one ortwo large LDs appeared in high-lipid cells (Fig. 1B). To maintainsimilar lipid contents and thus similar LDs between samples, weused cells cultured for 24 h in NL medium and for 48 h in PLmedium for further LD purification.

Zhu et al.

254 ec.asm.org March 2015 Volume 14 Number 3Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

LD purification and comprehensive proteomic analysis. Toget more insights into lipid accumulation, we purified LDs as wellas the lipids within LDs from R. toruloides cells according to apreviously described protocol (11, 26). It was found that the lipidsamples from the isolated LDs had significantly larger amounts ofneutral lipids but smaller amounts of polar lipids than the totallipid samples from the whole cells (Fig. 2A). Such a difference inlipid composition was a persuasive indication of the good purityof the isolated LDs. The protein profile of the LDs was uniquecompared with other cellular fractions (including total mem-brane, cytosol, and PNS), which further demonstrated that theisolated LDs were considerably pure (Fig. 2B). It was also clear thatthe purification procedure was reproducible because the proteinpatterns of LDs from three independent biological replicates werealmost identical (Fig. 2C and D).

To obtain an accurate LD proteome, a previously publishedmethod was applied (20). Briefly, the whole lane of YPD-24 h-3was cut into 29 slices (Fig. 2C, bands 1 to 29). In order to identifykey proteins involved in the maturation of large LDs and to checkthe comprehensiveness of the LD proteome, 7 visibly differentbands that appeared in LDs purified from the NL or PL sampleswere also collected (Fig. 2D, bands 30 to 36). The gel slices weredestained, and the proteins therein were digested according to thein-gel digestion procedure. The peptides were recovered and iden-

tified by LC-MS/MS. Thus, 224 proteins were identified from theYPD-24 h-3 sample, and two additional proteins were found inbands 31 and 34. Of these 226 proteins, 172 (76%) had functionalorthologs in S. cerevisiae (see Table S2 in the supplemental mate-rial). There were 63, 58, and 50 proteins whose orthologs have alsobeen found in the LD proteome of yeasts (S. cerevisiae, P. pastoris,and Y. lipolytica), humans (Homo sapiens), and fly (Drosophilamelanogaster), respectively (Fig. 3B), suggesting that at least 102(45%) were supported as LD proteins by those previous studies(13, 17, 18, 21, 28–30). These observations show that, for the firsttime, the LD proteome of the lipid-producing yeast R. toruloideswas established.

The LD proteome of R. toruloides was classified into 14 groupsaccording to the protein annotation and corresponding orthologsin S. cerevisiae (Fig. 3A; see also Table S2 in the supplementalmaterial). About one-third of the identified proteins were en-zymes involved in metabolism. There were 28 proteins associatedwith lipid metabolism (Table 1), including synthesis and degrada-tion of fatty acid, TAG, and sterol esters. Interestingly, most ofthem (20 proteins) were also found as LD proteins in other spe-cies. The presence of metabolic enzymes indicated that LD is notonly a passive reservoir but also an energetic organelle in whichmetabolic reactions occur. Other large groups included those in-volved in protein synthesis, folding, processing, degradation, andtransportation. There were 32 proteins that were identified as thecomponents of the ribosome or translational machinery and 10protein-folding-related chaperones, which were also frequentlyfound in the LDs of other species (21). For example, it has beendemonstrated that HSP70 of rat adipocytes is localized to LDsupon heat stimulation (37). Rab GTPases, coatomer components,v-SNARE, and ADP ribosylation factor are vesicle-trafficking-re-lated proteins, which play crucial roles in LD formation and sizedetermination. The fact that these proteins were found in LDs inthis study was consistent with previous observations (2, 7, 21). Wealso identified 34 mitochondrial proteins, which might be due tothe contamination that resulted from structural or functional in-teraction between LDs and mitochondria (38). A small propor-tion of LD proteins, such as two histones, were involved in geneticinformation storage and processing, which was consistent withthe results of a study on Drosophila (39). Moreover, 8 and 3 pro-teins were related to cellular signaling and cytoskeleton formation,respectively, and 21 proteins were poorly characterized in this LDproteome.

Interestingly, we found two potential LD structural proteins,RHTO_05627 and RHTO_03414. RHTO_05627 was previouslyannotated as a perilipin-like protein; here, we designated it Ldp1(lipid droplet protein 1). Fungal perilipin-like protein Mpl1 hasbeen first identified in Metarhizium anisopliae, and its homologsare present only in a small proportion of ascomycetous fungi, suchas the Pezizomycotina (40). However, at least one Ldp1 homologhas been found previously in basidiomycetous fungi (24). More-over, YALI0F24167g, the most abundant protein in the LD pro-teome of the ascomycetous yeast Y. lipolytica (13), was also thehomolog of Ldp1. RHTO_03414 is a caleosin family protein.Caleosins are LD-associated Ca2� binding proteins that have beenfound in many plants, several oleaginous fungi, and algae (3, 41).The biological functions of caleosins have been shown to stabilizethe oil body (alias of lipid droplet) of cycad megagametophytes(42) and to interact with vacuoles and promote degradation ofstorage lipids in Arabidopsis seeds (43). These results suggested

FIG 1 Lipid accumulation by R. toruloides cells. (A) Lipid contents of R.toruloides cells cultured under different conditions. Cells were collected fromcultures in YPD, nitrogen-limited (NL), and phosphorus-limited (PL) media.The total lipids were extracted and measured gravimetrically. The error barsrepresent the standard deviations for three independent samples. (B) Mor-phology of R. toruloides lipid droplets (LDs). Cells cultured in YPD mediumfor 24 h, NL medium for 24 h, or PL medium for 48 h were stained with NileRed and viewed by fluorescence microscope. BF, bright field microscopy; bar,10 �m.

Dynamics of R. toruloides Lipid Droplet Proteome

March 2015 Volume 14 Number 3 ec.asm.org 255Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

that LD structural proteins are likely conserved between diverseeukaryotic species.

The proteins identified from the bands with enhanced densi-tometry in the NL or PL samples are listed in Table 2. Besides the

proteins already found in the YPD-24 h-3 sample, RHTO_06504(conserved hypothetical protein) and RHTO_04226 (actin corti-cal patch component Lsb4, homolog of S. cerevisiae Lsb3) werenewly identified. The protein abundance of Ldp1, which migrated

FIG 2 Analysis of the lipids and proteins in purified lipid droplets (LDs) from R. toruloides. (A) Thin-layer chromatography analysis of lipid samples from wholecells (C) and LDs (L) prepared from cultures in YPD, nitrogen-limited (NL), or phosphorus-limited (PL) media. Solid lines and dotted lines on the left of thethree panels indicate lipid standards and unknown lipid species, respectively. T, triacylglycerol; F, fatty acid; D, diacylglycerol; M, monoacylglycerol; E,phosphatidylethanolamine; C, phosphatidylcholine. (B) SDS-PAGE analysis of proteins from LD, membrane, cytosol, and postnuclear supernatant (PNS)fractions. (C) SDS-PAGE analysis of LD proteins isolated from three independent samples cultured in YPD medium for 24 h. The whole lane of the YPD-24 h-3sample was cut into 29 slices and analyzed by LC-MS/MS, and the major proteins of each band are listed. (D) SDS-PAGE analysis of LD proteins isolated fromthree independent samples cultured in NL medium for 24 h or PL medium for 48 h and comparison of the protein patterns with those from a YPD culture. Theseven marked bands were cut for MS analysis, and the major proteins present in each band are listed on the right.

Zhu et al.

256 ec.asm.org March 2015 Volume 14 Number 3Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

to the bands identical to band 31, was different in these samples.The abundance of Ldp1 was highest in the NL sample while lowestin the YPD sample (Fig. 2D), suggesting that it might be involvedin lipid overproduction and giant LD formation. RHTO_07686, a123-kDa protein similar to Aspergillus nidulans psi factor-produc-ing oxygenase PpoA, was also identified. PpoA is a bifunctionalP450 protein that oxidizes linoleic acid to (8R)-8-hydroper-oxyoctadecadienoic acid followed by the isomerization to form(5S,8R)-5,8-dihydroxyoctadecadienoic acid (44). The corre-sponding oxylipin products are hormone-like signaling mediators

regulating the balance between asexual and sexual life cycles (44,45). It has been demonstrated that PpoA is localized to the LDs inA. nidulans (45), but detailed roles of PpoA in LD function andlipid accumulation remained unclear. We also found a higher-molecular-mass PpoA isoform (about 170 kDa) whose abundancewas higher in the YPD and NL samples than in the PL sample (Fig.2D). RHTO_03117, one of the two adenylate kinases and the or-tholog of S. cerevisiae cytosolic Adk1, was the major protein ofband 36. It was remarkable to find RHTO_03117 in the LD pro-teome obtained from the PL sample. Under PL conditions, the

FIG 3 Function catalogs of the R. toruloides lipid droplet (LD) proteome. (A) Functional classification of the identified proteins. (B) Venn diagram of proteinspresent in the R. toruloides LD proteome that have homologs in yeasts (S. cerevisiae, P. pastoris, and Y. lipolytica), humans (Homo sapiens), and fly (Drosophilamelanogaster). The function of each protein is listed in Table S2 in the supplemental material.

TABLE 1 Identified LD proteins involved in lipid metabolism

Function catalog(total no. of proteins) Identified proteinsa

Fatty acid synthesis (4) RHTO_03915 (ATP citrate synthase), RHTO_02004 (acetyl-CoA carboxylase, Acc1), RHTO_02032 (fatty acid synthase subunitbeta, fungus type, Fas1), RHTO_02139 (fatty acid synthase subunit alpha, fungus type, Fas2)

TAG synthesis (3) RHTO_03193 (1-acylglycerone phosphate reductase, Ayr1), RHTO_07665 (glycerol-3-phosphate dehydrogenase, Gut2),RHTO_04314 (glycerol-3-phosphate/dihydroxyacetone phosphate dual substrate-specific sn-1 acyltransferase, Gpt2)

Steroid synthesis (4) RHTO_00856 (delta(24)-sterol C-methyltransferase, Erg6), RHTO_01745 (squalene monooxygenase, Erg1), RHTO_02048(acetyl-CoA C-acetyltransferase, Erg10), RHTO_04192 (lanosterol synthase, Erg7)

Hydrolase (9) RHTO_01062 (steryl ester hydrolase, Yeh2), RHTO_01389 (lipase, class 3), RHTO_03165 (inositol or phosphatidylinositolphosphatase, Inp51), RHTO_03511 (alpha/beta hydrolase fold protein, Yju3), RHTO_03913 (monoglyceride lipase,homolog of YJU3, Yju3), RHTO_05588 (N-acyl-phosphatidylethanolamine-hydrolyzing phospholipase D, Fmp30),RHTO_03931 (bifunctional enzyme with triacylglycerol lipase and lysophosphatidic acid acyltransferase activity, Tgl5),RHTO_03669 (esterase/lipase/thioesterase family protein), RHTO_05515 (alpha/beta hydrolase, putative lipase/esterase)

Fatty acid oxidation (5) RHTO_00397 (acyl-CoA dehydrogenase), RHTO_01116 (acyl-CoA thioesterase 8, Tes1), RHTO_03348 (3-oxoacyl-[acyl-carrier protein] reductase), RHTO_07686 (fatty acid oxygenase PpoA), RHTO_05680 (putative fatty aldehydedehydrogenase, Hfd1)

Others (3) RHTO_07994 (phosphatidylserine decarboxylase, Psd1), RHTO_04135 (lipase/thioesterase family protein, similar to steroldeacetylase Say1), RHTO_03216 (putative acyltransferase with similarity to Eeb1p and Eht1p, YMR210W)

a The names of corresponding orthologs in S. cerevisiae are underlined.

Dynamics of R. toruloides Lipid Droplet Proteome

March 2015 Volume 14 Number 3 ec.asm.org 257Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

cellular levels of free phosphate and ATP and the energy chargemay be low, and adenylate kinase is responsible for converting twoADP molecules into one ATP and one AMP to maintain ATPhomeostasis (46). RHTO_04363, another abundant protein iden-tified in the PL sample, was a mitochondrial L-malate dehydroge-nase.

Expression dynamics of the LD proteome. The results showeda dramatic increase in lipid content when the cells were culturedunder NL and PL conditions (Fig. 1), indicating that R. toruloidesis a good model to study LD dynamics and identify the key pro-teins involved in the formation of large LDs. The SDS-PAGE re-sults indicated that the LD proteome changed substantially. Themost significant change was an enhanced densitometry of bands31 and 32 in the NL sample and of band 36 in the PL samplecompared with those in the YPD sample (Fig. 2D). However, thisassay was inadequate to offer more quantitative information. Thedynamic changes of the LD proteins might have resulted from theregulation of gene expression and organelle targeting. The infor-mation about expression changes of LD-related genes should beincluded in the global differential transcriptome and proteomedata sets. In our previous work, we have sequenced the transcript3=-end tags and performed a quantitative comparison of the tran-scriptomes between a sample cultured in minimal medium (MM)containing 47 mM NH4

� and a sample in nitrogen-limited min-imal medium (MM-N). The lipid contents of the MM and theMM-N samples were 22.8% and 33.3%, respectively, and the ex-pression levels of over 2,000 genes were significantly different inthese two samples (24). Interestingly, in this study we found thatmore than one-half of the LD-associated genes were expressedwithin a 2-fold variation between the MM and MM-N samplesand only 9 genes were not detected in the 3=-tag digital gene ex-pression profiling analysis (Fig. 4A). Of those with a �2-foldchange, 35 and 50 genes were up- and downregulated, respec-tively, in the MM-N sample (Fig. 4A; see also Tables S3 and S4 inthe supplemental material). We also compared the proteomes ofcells from the YPD culture (“Seed”), the NL culture for 24 h (“24h”), and the NL culture for 96 h (“96 h”) (24, 25). Similarly, thelipid contents increased from 9.8% in the “Seed” sample to 63% inthe “96 h” sample. The levels of over 500 proteins significantlychanged among these samples (24), of which 90% of the LD-associated proteins (203 proteins) were identified in at least onesample (Fig. 4B). By rigorous statistical testing, the proteins that

significantly changed in the NL samples in comparison with thosein the nutrient-rich sample were found (Fig. 4B). Protein abun-dances changed similarly in the NL sample over time, because theproportion of overlap between the 24 h/Seed data set and the 96h/Seed data set was high (Fig. 4C). The changed proteins in the “24h” and “96 h” samples were merged, and 38 and 39 proteins wereup- and downregulated, respectively, under nitrogen-limited con-ditions (see Tables S3 and S4 in the supplemental material).

The reanalysis of the previous comparative transcriptome andproteome data clearly revealed that the transcription and expres-sion levels of many LD-related genes changed dynamically duringlipid accumulation. In the current work, nitrogen or phosphoruslimitation also altered the protein profiles of the LD proteome(Fig. 2D). In order to examine whether the expression of the LD-associated genes was regulated by nutrient limitation, RT-qPCRwas performed to analyze the transcriptional levels of 9 selectedgenes (Fig. 4D). These genes encoded proteins important for LDmaintenance, enlargement, and mobilization. It was found that 6genes were transcriptionally upregulated, which was coincidentwith the “omic” analysis. The protein abundance of Ldp1 in-creased more than 200-fold in the “96 h” sample compared withthat in the “Seed” sample (Fig. 4D). The RT-qPCR analysis con-firmed the upregulation of LDP1 in the MM-N sample (data notshown), although the transcriptome analysis failed to detect theexpression level of LDP1, most likely because of the absence of theCATG site to generate the transcript tag. The RT-qPCR resultsalso clearly showed that the transcriptional level of LDP1 was up-regulated under both NL and PL conditions. Moreover, the tran-scription level was higher in the NL sample than in the PL sample(Fig. 4D). Interestingly, protein levels of Acl1 (ATP:citrate lyase)and Acc1 (acetyl-CoA carboxylase), key enzymes for the forma-tion of precursors for lipid biosynthesis (24), were upregulatedunder the NL condition. The RT-qPCR analysis indicated that thetranscription of ACL1 and ACC1 was elevated in both the NL-24 hand the PL-48 h samples and that the transcription levels in the PLsample were lower than that in the NL sample (Fig. 4D). TheRT-qPCR results showed that the transcription of FAS1 (fatty acidsynthase, beta subunit) was upregulated about 8-fold under NLand PL conditions, which was also in agreement with previoustranscriptome and proteome results. Under nutrient-limited con-ditions, higher protein levels of PpoA, the enzyme for oxylipinproduction, were also confirmed by RT-qPCR. Although the tran-

TABLE 2 Identified proteins from bands 30 to 36

SDS-PAGE band(total no. ofproteins in thecatalog) Identified proteinsa

Band 30 (1) RHTO_07686 (fatty acid oxygenase PpoA)Band 31 (3) LDP1 (perilipin-like), RHTO_06504 (conserved hypothetical protein), RHTO_07648 (voltage-dependent ion-selective channel, Por1)Band 32 (2) RHTO_07648 (voltage-dependent ion-selective channel, Por1), LDP1 (perilipin-like)Band 33 (1) RHTO_07686 (fatty acid oxygenase PpoA)Band 34 (3) RHTO_04363 (L-malate dehydrogenase, mitochondrial, Mdh1), RHTO_00208 (short-chain dehydrogenase/reductase SDR family

protein, YOR246C), RHTO_00474 (mitochondrial phosphate carrier protein, Pic2)Band 35 (6) RHTO_04363 (L-malate dehydrogenase, mitochondrial, Mdh1), RHTO_03117 (adenylate kinase, Adk1), RHTO_06575 (NmrA-like

domain containing protein), RHTO_03193 (1-acylglycerone phosphate reductase, Ayr1), RHTO_06960 (protein of glucose/ribitoldehydrogenase family, weak similarity to YKL107W), RHTO_04226 (actin cortical patch component Lsb4, Lsb3),

Band 36 (2) RHTO_03117 (adenylate kinase, Adk1), LDP1 (perilipin-like)a Proteins in bold are the major proteins for which the most peptides were identified. The names of corresponding orthologs in S. cerevisiae are underlined.

Zhu et al.

258 ec.asm.org March 2015 Volume 14 Number 3Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

scription of 4 genes, RHTO_00856 and RHTO_04192 for steroidsynthesis and RHTO_01062 and RHTO_03165 for lipid hydroly-sis, were all upregulated in the MM-N sample, RT-qPCR resultsindicated that only RHTO_01062 was significantly upregulatedunder NL conditions. The other three were downregulated underNL or PL conditions (Fig. 4D).

In the LD proteome of R. toruloides, several other lipid meta-bolic enzymes, such as acyl coenzyme A (acyl-CoA) dehydroge-nase (RHTO_00397), lipase (RHTO_01389 and RHTO_03913),and fatty aldehyde dehydrogenase (RHTO_05680), were upregu-lated in the NL samples, while glycerol-3-phosphate dehydroge-nase (RHTO_007665), acyl-CoA thioesterase 8 (RHTO_01116),and lipid hydrolase (RHTO_03669) were downregulated. Thelipid biosynthesis-associated proteins, such as 3-oxoacyl-ACP re-

ductase (RHTO_03348) and acetyl-CoA C-acetyltransferase(RHTO_02048), were transcriptionally upregulated in the MM-Nsample, but the latter was downregulated in the NL sample (seeTable S3 in the supplemental material).

Ldp1 localized on LDs and promoted giant LD formation.Since Ldp1 has a putative perilipin domain (Pfam numberPF03036) and it is a potential LD structural protein, we carried outexperiments to verify its LD localization, to examine its cellulardistribution, and more importantly, to determine whether it was akey player for giant LD formation. First, we made a polyclonalantibody specifically recognizing Ldp1. To determine the cellulardistribution of Ldp1, we fractionated cell lysates into LDs, totalmembrane, cytosol, and PNS during the LD purification process.Proteins separated by SDS-PAGE were immunoblotted with the

FIG 4 Dynamics of lipid droplet proteome. (A) Relative transcription levels of LD-associated genes revealed from digital gene expression analysis of the cellscultured in minimum medium (MM) and nitrogen-limited minimum medium (MM-N). The MM sample was used as a reference. (B) Relative protein levels ofLD proteins revealed from comparative proteome analysis. Protein levels of the cells cultured in nutrient-rich medium (“Seed”) were used as a reference, andrelative protein levels and differentially expressed proteins in cells cultured in nitrogen-limited (NL) medium (“24 h” and “96 h”) are shown. (C) Overlap of thedifferentially expressed LD proteins in the “24 h” and “96 h” samples. (D) Relative mRNA expression levels between cells cultured in nutrient-limited medium(“NL-24 h” and “PL-48 h”) and nutrient-rich medium (“YPD-24 h”) revealed by RT-qPCR. Three biological replicates and two technical replicates wereperformed. Statistical significance (*, P 0.05; **, P 0.01; ***, P 0.001) was determined by one-tailed Student’s t test. The error bars represent the standarderrors for three independently cultured samples. The expression levels were normalized to two reference genes (RHTO_03560 and RHTO_03746). The geneannotation and the transcript and protein expression levels, respectively, from comparative transcriptome (panel A) and proteome (panel B) analysis are listedin the right panel. Normalized counts of transcript tags (TPM, tags per million) were used for comparison of transcription levels. The average of the normalizedspectral counts from three replicates was used for comparison of the protein levels. The comparative transcriptome and proteome data were retrieved from ourprevious study (24).

Dynamics of R. toruloides Lipid Droplet Proteome

March 2015 Volume 14 Number 3 ec.asm.org 259Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

anti-Ldp1 antibody, and the results clearly showed that Ldp1 waspresent only in the LD fraction, even with much lower proteinloading than in other cellular fractions (Fig. 5A). These data con-firmed the LD localization of Ldp1 and suggested Ldp1 as an LD-specific protein in R. toruloides. We then compared the proteinabundance of Ldp1 in the LD fractions prepared from cells cul-tured under different conditions and found that the protein levelsin both the NL-24 h and PL-48 h samples were higher than that in

the YPD-24 h sample. Moreover, the NL-24 h sample had a higherlevel of Ldp1 than the PL-48 h sample (Fig. 5B), which matchedwell with the densitometry change of the protein bands that mi-grated identically to band 31 between these samples (Fig. 2D).These results were also in agreement with results from semiquan-titative proteome and RT-qPCR analysis (Fig. 4D). Together,these observations clearly demonstrated the dynamic nature ofLdp1 during lipid production. Second, we expressed a C-terminal

FIG 5 Localization and function of the lipid droplet (LD) structure protein Ldp1. (A) Distribution of Ldp1 in different cellular fractions. The proteins from LDs(LD), total membrane (Membrane), cytosol (Cytosol), and postnuclear supernatant (PNS) were separated by SDS-PAGE and blotted with anti-Ldp1 antibody.(B) Comparison of Ldp1 protein levels in the LD proteome under different culture conditions. Equal amounts of LD proteins from R. toruloides cells cultured inYPD for 24 h, NL for 24 h, and PL for 48 h were separated by SDS-PAGE and blotted by anti-Ldp1 antibody. The arrows in the left panels indicate the positionof Ldp1, and the right panels were colloidal blue stained and used as loading control (A and B). (C) Localization of heterogeneously expressed Ldp1 in S. cerevisiae.The localizations of Ldp1-GFP, Erg6-RFP (LD marker), and Sec13-RFP (the marker for ER to Golgi vesicles) were compared. (D) Overexpressed Ldp1-GFPlocalized on LDs and facilitated the formation of giant LDs in S. cerevisiae. S. cerevisiae cells expressing GFP or Ldp1-GFP were observed by confocal microscope.LDs were stained with LipidTOX. Bar, 5 �m (C and D).

Zhu et al.

260 ec.asm.org March 2015 Volume 14 Number 3Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

GFP-tagged Ldp1 (Ldp1-GFP) in S. cerevisiae lacking perilipin-like proteins to morphologically determine the cellular distribu-tion of Ldp1. The Ldp1-GFP colocalized well with the LD markerErg6-RFP (47) and LipidTOX staining but not with Sec13-RFP,which is a marker protein for endoplasmic reticulum (ER) toGolgi vesicles (Fig. 5C and D), while the expressed GFP distrib-uted in the cytosol space (d1 to d5 in Fig. 5D). We also examinedthe LD sizes of S. cerevisiae cells overexpressing Ldp1-GFP. Inter-estingly, most cells that expressed Ldp1-GFP contained one giantLD, whereas the cells that expressed GFP had multiple small LDs(Fig. 5D; see also Fig. S1 in the supplemental material), indicatingthat the overexpression of Ldp1 was responsible for LD enlarge-ment.

DISCUSSION

R. toruloides is a basidiomycete fungus of the Pucciniomycotinasubphylum. Many species of the Pucciniomycotina clade are plant-parasitic rust fungi that can cause severe forest and agriculturelosses (48). It has been known that LDs are connected with patho-gen virulence of fungi (40, 49). However, there have been no sys-tematic studies on the LD proteome of basidiomycetes. Moreover,this yeast is well known for the overproduction of lipids and pig-ments (15, 22, 23, 50) and has been used as a microbial factory forthe production of advanced biofuels (15, 51). Manipulating thedynamics of LDs is a potential approach to engineer this yeast forthe production of lipids and related metabolites at higher titer,rate, and yield. For example, the ER proteins seipin and fat stor-age-inducing transmembrane proteins (FIT1 and FIT2) have beenfound to regulate LD formation and morphology (52), and im-pairing the expression of seipin can increase TAG synthesis andlead to giant LD formation (53, 54). In addition, higher levels ofFIT1 or FIT2 have resulted in more and larger LDs (55). Theexpression of the LD structure protein caleosin 1 (AtClo1) fromArabidopsis in S. cerevisiae increased lipid accumulation (56). Re-cently, many efforts have been undertaken to engineer S. cerevisiaeor Escherichia coli for the production of lipid-derived metabolitesand biofuels, but the efficiency remained fairly low (14, 57). Alter-natively, it has been shown that more than 70% of the TAGs in R.toruloides can be converted to fatty acid ethyl esters when incubat-ing lipid-rich cells with 10% ethanol in an aqueous environment(58). It is believed that the LD provides the hydrophobic microen-vironment for LD-associated lipases, which traditionally catalyzedthe transesterification in nonaqueous media (59). Although theroles of LDs are important in lipid biotechnology, key factors reg-ulating the LD dynamics remain poorly characterized in oleagi-nous species.

In this study, we established the LD proteome of R. toruloides.By analyzing the protein profiles of the LDs from high-lipid andlow-lipid cell samples, we found that the LD proteome changedsignificantly (Fig. 2D). We speculated that these proteomechanges were crucial for lipid accumulation and LD dynamics;thus, we reanalyzed the data sets of our previous comparativetranscriptome and proteome study to identify differentially ex-pressed LD-associated genes, as those previous samples were cul-tured in nitrogen-rich and nitrogen-limited media and accumu-lated different amounts of lipids (24, 25). The expression levels ofabout one-third of the LD-associated genes changed (see Table S3in the supplemental material), while only 25% of the overall genesand 7% of the whole proteome of the cell changed at the transcrip-

tome and proteome levels, respectively. This indicated that the LDproteome changed more significantly than other cellular proteins.

The LDs are hubs of cellular lipid metabolism and can serve asthe reservoir of energy or building blocks of cellular membranelipids. Acetyl-CoA, a precursor for fatty acid synthesis, is gener-ated from the cleavage of citrate, which permeates out from themitochondrion in oleaginous yeasts (22). The enzyme catalyzingcitrate to acetyl-CoA is Acl1. Acetyl-CoA is activated to malonyl-CoA by acetyl-CoA carboxylase (Acc), and then malonyl-CoA isutilized for fatty acid chain elongation, which is catalyzed by fattyacid synthase (Fas), a multifunctional enzyme complex. Thesethree enzymes were traditionally known as cytoplasmic proteins,but they were found in the LD proteome in this study as well asothers (60). It was unknown whether these proteins were translo-cated to LDs by vesicle transport or were recruited from the cyto-sol. Other lipid biosynthesis-related proteins were found in theLD proteome, including RHTO_07665, RHTO_03193, andRHTO_04314. All these enzymes constituted a complete pathwayfrom citrate and dihydroxyacetone phosphate to lysophosphatidicacid, a common precursor for TAGs and phospholipids. Althoughthe enzymes responsible for the downstream biosynthetic steps ofTAGs and phospholipids, such as Slc1 and Dga1, were found inthe LD proteomes of mammalian and yeasts (13, 17, 21, 38, 61),they were not identified in the LD proteome of R. toruloides. Itshould be noted that a cytosol TAG synthesis machinery has beenknown in the oleaginous red yeasts (62), but its roles in lipid ac-cumulation and TAG transport remained unclear. Of these LD-associated lipid biosynthesis genes, ACL1, ACC1, FAS1, and FAS2were all upregulated in lipid-rich cells at the protein level (seeTable S3 in the supplemental material) as well as the transcrip-tional level (Fig. 4D).

The LDs are the major organelles for neutral lipid mobilizationin R. toruloides. We found 11 lipolytic enzymes in the LD pro-teome (Table 1). RHTO_03931 is the ortholog of the yeast bifunc-tional enzyme Tgl5, which has TAG lipase and lysophosphatidicacid acyltransferase activity. This enzyme could transfer the acylgroup from acyl-CoA to lysophosphatidic acid to produce phos-phatidic acid, which could promote LD fusion to generate largerLDs (52, 53). This enzyme could also hydrolyze TAG to fatty acidand diacylglycerol, which could then be hydrolyzed to monoacyl-glycerol by RHTO_02359 (the ortholog of mammalian hormone-sensitive lipase). The monoacylglycerol was hydrolyzed to fattyacid and glycerol by lipases RHTO_03913 and probably RHTO_03511. However, RHTO_02359 was absent in the LD proteome,suggesting that it was not recruited yet at the lipid accumulationstage, because the translocation of the activated and phosphory-lated hormone-sensitive lipase onto LDs depends on phosphory-lation of the perilipin (Ldp1 might play similar roles in this yeast)by protein kinase A in response to the cellular energy status (19).Released fatty acids may be metabolized, such as the transforma-tion of linoleic acid to oxylipin by PpoA (RHTO_07686). Alter-natively, fatty acids may be oxidized via the -oxidation process.The location of -oxidization is believed to be in the mitochon-drion and peroxisome, which are both in close proximity to LDs.Indeed, mitochondrial and peroxisomal proteins (RHTO_01116,RHTO_03348, and RHTO_00397) were identified in the LD pro-teome. PpoA was upregulated in lipid-rich cells (Fig. 4D). Inter-estingly, the expression levels of three genes for lipid degradation(RHTO_03913, RHTO_01116, and RHTO_03348) were up-

Dynamics of R. toruloides Lipid Droplet Proteome

March 2015 Volume 14 Number 3 ec.asm.org 261Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

regulated in the MM-N sample (see Table S3 in the supplementalmaterial), in which the -oxidation was elevated (24).

Membrane trafficking is a delivery process mediated by multi-organelles. The expression patterns of proteins involved in mem-brane trafficking can influence LD dynamics. RHTO_07779, anADP-ribosylation factor (ARF1) and a homolog of DrosophilaARF79F, was found in the LD proteome. ARF1 is a small GTPase,which recruits the COPI coatomer subunits and mediates Golgiapparatus-to-ER transport. Knockdown of ARF79F and COPIcomponents has been reported to promote lipid accumulation,partially due to the decrease of the LD targeting of LD lipase ATGL(63, 64). RHTO_07779 was downregulated in the “96 h” sample,which had a lipid content of 63%. Three COPI components, sub-units �, , and � corresponding to RHTO_00667, RHTO_00612,and RHTO_03548, respectively, were also found in the LD pro-teome, but the expression of the subunit was upregulated in thelipid-rich samples (see Table S3 in the supplemental material). Itwas also known that the COPII controlled by small GTPase SAR1was involved in protein anterograde transport from the ER to theGolgi apparatus and played similar roles in ATGL translocation toLDs (65). Impaired COPII activity increased lipid storage but ina milder manner. We identified Sar1 GTPase (RHTO_02448)in the LD proteome. This protein was downregulated in the “24h” and “96 h” samples, but its transcription level increased2.6-fold in the MM-N sample. Two vesicle fusion-related proteins,RHTO_01157 (Ykt6, v-SNARE) and RHTO_06548 (Sec18, an ortholog tohuman N-ethylmaleimide-sensitive factor [NSF]) (66), were alsopresent in the LD proteome.

Small Rab GTPases are vesicle-trafficking-regulating G pro-teins with posttranslationally modified prenyl moieties for mem-brane anchorage. Seven Rabs were found in the LD proteome,probably due to the presence of ER-derived phospholipidhemimembrane of LDs. Rabs may mediate the interactions be-tween LDs and other organelles, such as the Golgi apparatus, vac-uoles, and endosomes (2, 67). The expression of small Rabsalso changed during lipid overproduction, such as upregulationof Rab1 (RHTO_06528), Rab8 (RHTO_08047), and Rab11(RHTO_01102) in the NL sample (see Table S3 in the supplemen-tal material). RHTO_04520, a homolog to Rab18, was upregu-lated in the MM-N sample. Interestingly, a higher upregulatedexpression level of Rab18 was found in fatty adipocytes, and re-cruitment of Rab18 to LDs in association with lipolysis was dem-onstrated (68). Thus, RHTO_04520 may play a role in LD metab-olism. Other Rabs (Rab4, -5, -7, and -11) may be associated withendosomes or vacuoles. Vacuoles are organelles playing a majorrole in autophagy, the process of “eating” cellular components. Itwas also demonstrated that autophagy played crucial roles in theformation of giant LDs in adipocytes (69). In our previous study,an elevated autophagy process, which might remodel the cells tomake more space for LDs, was revealed along with lipid accumu-lation (24).

Taken together, this and many recent studies clearly indicatethat LD-associated proteins are not simply sticky but structurallyand functionally associated with LDs. The presence of metabolicenzymes (involved mainly in lipid metabolism), vesical trafficproteins, Rab GTPases, and others in the LD proteome of differentspecies suggests more-complicated and cryptic roles of these pro-teins in LD biology. The presence of functionally diverse proteinsalso suggests that the cytosolic LDs are not simple energy storage

particles but more energetic and functional organelles. More im-portantly, our results showed that the levels of the majority of theLD-associated proteins changed dynamically in response to nutri-ent availability. Such changes had good correlation with the de-velopment of cellular LDs.

In summary, we identified 226 LD-associated proteins, whichestablished a comprehensive LD proteome of the oleaginous yeastR. toruloides. The major LD structure protein Ldp1 was character-ized as an LD marker protein, which was important for giant LDformation. Analysis of proteomic data and biochemical resultsdemonstrated the dynamic nature of these proteins during lipidaccumulation under different conditions. Our results significantlyadvance our understanding of the molecular basis of lipid over-production in oleaginous yeasts. Such information will be valu-able for the investigation on LD biology in general and for thedevelopment of superior lipid producers.

ACKNOWLEDGMENTS

We thank Hongyuan Yang for his good suggestions on LD morphology,Erin O’Shea for sharing the strains expressing Erg6-RFP and Sec13-RFP,and Nicolaas A. A. Buijs for critically reading the manuscript.

This work was funded by the Natural Sciences Foundation of China(31370128, 31170060, and 21325627) and the National Basic Researchand Development Program of China (2011CB707405 and 2011CBA00906).

REFERENCES1. Wältermann M, Steinbüchel A. 2005. Neutral lipid bodies in pro-

karyotes: recent insights into structure, formation, and relationship toeukaryotic lipid depots. J Bacteriol 187:3607–3619. http://dx.doi.org/10.1128/JB.187.11.3607-3619.2005.

2. Walther TC, Farese RV, Jr. 2012. Lipid droplets and cellular lipidmetabolism. Annu Rev Biochem 81:687–714. http://dx.doi.org/10.1146/annurev-biochem-061009-102430.

3. Murphy DJ. 2001. The biogenesis and functions of lipid bodies in ani-mals, plants and microorganisms. Prog Lipid Res 40:325– 438. http://dx.doi.org/10.1016/S0163-7827(01)00013-3.

4. Martin S, Parton RG. 2006. Lipid droplets: a unified view of a dynamicorganelle. Nat Rev Mol Cell Biol 7:373–378. http://dx.doi.org/10.1038/nrm1912.

5. Fujimoto T, Ohsaki Y, Cheng J, Suzuki M, Shinohara Y. 2008. Lipiddroplets: a classic organelle with new outfits. Histochem Cell Biol 130:263–279. http://dx.doi.org/10.1007/s00418-008-0449-0.

6. Farese RV, Jr, Walther TC. 2009. Lipid droplets finally get a little R-E-S-P-E-C-T. Cell 139:855– 860. http://dx.doi.org/10.1016/j.cell.2009.11.005.

7. Saka HA, Valdivia R. 2012. Emerging roles for lipid droplets in immunityand host-pathogen interactions. Annu Rev Cell Dev Biol 28:411– 437.http://dx.doi.org/10.1146/annurev-cellbio-092910-153958.

8. Singh R, Kaushik S, Wang Y, Xiang Y, Novak I, Komatsu M, Tanaka K,Cuervo AM, Czaja MJ. 2009. Autophagy regulates lipid metabolism.Nature 458:1131–1135. http://dx.doi.org/10.1038/nature07976.

9. Zehmer JK, Huang Y, Peng G, Pu J, Anderson RG, Liu P. 2009. A rolefor lipid droplets in inter-membrane lipid traffic. Proteomics 9:914 –921.http://dx.doi.org/10.1002/pmic.200800584.

10. Yang L, Ding Y, Chen Y, Zhang S, Huo C, Wang Y, Yu J, Zhang P, NaH, Zhang H, Ma Y, Liu P. 2012. The proteomics of lipid droplets:structure, dynamics, and functions of the organelle conserved from bac-teria to humans. J Lipid Res 53:1245–1253. http://dx.doi.org/10.1194/jlr.R024117.

11. Ding Y, Yang L, Zhang S, Wang Y, Du Y, Pu J, Peng G, Chen Y, ZhangH, Yu J, Hang H, Wu P, Yang F, Yang H, Steinbuchel A, Liu P. 2012.Identification of the major functional proteins of prokaryotic lipid drop-lets. J Lipid Res 53:399 – 411. http://dx.doi.org/10.1194/jlr.M021899.

12. Moellering ER, Benning C. 2010. RNA interference silencing of a majorlipid droplet protein affects lipid droplet size in Chlamydomonas rein-hardtii. Eukaryot Cell 9:97–106. http://dx.doi.org/10.1128/EC.00203-09.

13. Athenstaedt K, Jolivet P, Boulard C, Zivy M, Negroni L, Nicaud JM,Chardot T. 2006. Lipid particle composition of the yeast Yarrowia lipoly-

Zhu et al.

262 ec.asm.org March 2015 Volume 14 Number 3Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

tica depends on the carbon source. Proteomics 6:1450 –1459. http://dx.doi.org/10.1002/pmic.200500339.

14. Steen EJ, Kang Y, Bokinsky G, Hu Z, Schirmer A, McClure A, DelCardayre SB, Keasling JD. 2010. Microbial production of fatty-acid-derived fuels and chemicals from plant biomass. Nature 463:559 –562.http://dx.doi.org/10.1038/nature08721.

15. Li Y, Zhao Z, Bai F. 2007. High-density cultivation of oleaginous yeastRhodosporidium toruloides Y4 in fed-batch culture. Enzyme Microb Tech-nol 41:312–317. http://dx.doi.org/10.1016/j.enzmictec.2007.02.008.

16. Bartz R, Li WH, Venables B, Zehmer JK, Roth MR, Welti R, AndersonRG, Liu P, Chapman KD. 2007. Lipidomics reveals that adiposomes storeether lipids and mediate phospholipid traffic. J Lipid Res 48:837– 847.http://dx.doi.org/10.1194/jlr.M600413-JLR200.

17. Grillitsch K, Connerth M, Kofeler H, Arrey TN, Rietschel B, Wagner B,Karas M, Daum G. 2011. Lipid particles/droplets of the yeast Saccharo-myces cerevisiae revisited: lipidome meets proteome. Biochim BiophysActa 1811:1165–1176. http://dx.doi.org/10.1016/j.bbalip.2011.07.015.

18. Ivashov VA, Grillitsch K, Koefeler H, Leitner E, Baeumlisberger D,Karas M, Daum G. 2013. Lipidome and proteome of lipid droplets fromthe methylotrophic yeast Pichia pastoris. Biochim Biophys Acta 1831:282–290. http://dx.doi.org/10.1016/j.bbalip.2012.09.017.

19. Bickel PE, Tansey JT, Welte MA. 2009. PAT proteins, an ancient familyof lipid droplet proteins that regulate cellular lipid stores. Biochim Bio-phys Acta 1791:419 – 440. http://dx.doi.org/10.1016/j.bbalip.2009.04.002.

20. Bartz R, Zehmer JK, Zhu M, Chen Y, Serrero G, Zhao Y, Liu P. 2007.Dynamic activity of lipid droplets: protein phosphorylation and GTP-mediated protein translocation. J Proteome Res 6:3256 –3265. http://dx.doi.org/10.1021/pr070158j.

21. Hodges BD, Wu CC. 2010. Proteomic insights into an expanded cellularrole for cytoplasmic lipid droplets. J Lipid Res 51:262–273. http://dx.doi.org/10.1194/jlr.R003582.

22. Ratledge C, Wynn JP. 2002. The biochemistry and molecular biology oflipid accumulation in oleaginous microorganisms. Adv Appl Microbiol51:1–51. http://dx.doi.org/10.1016/S0065-2164(02)51000-5.

23. Wu S, Hu C, Jin G, Zhao X, Zhao ZK. 2010. Phosphate-limitationmediated lipid production by Rhodosporidium toruloides. Bioresour Tech-nol 101:6124 – 6129. http://dx.doi.org/10.1016/j.biortech.2010.02.111.

24. Zhu Z, Zhang S, Liu H, Shen H, Lin X, Yang F, Zhou YJ, Jin G, Ye M,Zou H, Zhao ZK. 2012. A multi-omic map of the lipid-producing yeastRhodosporidium toruloides. Nat Commun 3:1112. http://dx.doi.org/10.1038/ncomms2112.

25. Liu H, Zhao X, Wang F, Li Y, Jiang X, Ye M, Zhao ZK, Zou H. 2009.Comparative proteomic analysis of Rhodosporidium toruloides during lipidaccumulation. Yeast 26:553–566. http://dx.doi.org/10.1002/yea.1706.

26. Ding Y, Zhang S, Yang L, Na H, Zhang P, Zhang H, Wang Y, Chen Y,Yu J, Huo C, Xu S, Garaiova M, Cong Y, Liu P. 2013. Isolating lipiddroplets from multiple species. Nat Protoc 8:43–51. http://dx.doi.org/10.1038/nprot.2012.142.

27. Zhao X, Kong X, Hua Y, Feng B, Zhao ZK. 2008. Medium optimizationfor lipid production through co-fermentation of glucose and xylose by theoleaginous yeast Lipomyces starkeyi. Eur J Lipid Sci Technol 110:405– 412.http://dx.doi.org/10.1002/ejlt.200700224.

28. Su W, Wang Y, Jia X, Wu W, Li L, Tian X, Li S, Wang C, Xu H, CaoJ, Han Q, Xu S, Chen Y, Zhong Y, Zhang X, Liu P, Gustafsson JA, GuanY. 2014. Comparative proteomic study reveals 17beta-HSD13 as a patho-genic protein in nonalcoholic fatty liver disease. Proc Natl Acad Sci U S A111:11437–11442. http://dx.doi.org/10.1073/pnas.1410741111.

29. Krahmer N, Hilger M, Kory N, Wilfling F, Stoehr G, Mann M, FareseRV, Walther TC. 2013. Protein correlation profiles identify lipid dropletproteins with high confidence. Mol Cell Proteomics 12:1115–1126. http://dx.doi.org/10.1074/mcp.M112.020230.

30. Currie E, Guo X, Christiano R, Chitraju C, Kory N, Harrison K, HaasJ, Walther TC, Farese RV. 2014. High confidence proteomic analysis ofyeast LDs identifies additional droplet proteins and reveals connections todolichol synthesis and sterol acetylation. J Lipid Res 55:1465–1477. http://dx.doi.org/10.1194/jlr.M050229.

31. Jiang X, Jiang X, Han G, Ye M, Zou H. 2007. Optimization of filteringcriterion for SEQUEST database searching to improve proteome coveragein shotgun proteomics. BMC Bioinformatics 8:323. http://dx.doi.org/10.1186/1471-2105-8-323.

32. Wan J, Roth AF, Bailey AO, Davis NG. 2007. Palmitoylated proteins:purification and identification. Nat Protoc 2:1573–1584. http://dx.doi.org/10.1038/nprot.2007.225.

33. Vandesompele J, De Preter K, Pattyn F, Poppe B, Van Roy N, De PaepeA, Speleman F. 2002. Accurate normalization of real-time quantitativeRT-PCR data by geometric averaging of multiple internal control genes.Genome Biol 3:RESEARCH0034. http://dx.doi.org/10.1186/gb-2002-3-7-research0034.

34. Heckman KL, Pease LR. 2007. Gene splicing and mutagenesis by PCR-driven overlap extension. Nat Protoc 2:924 –932. http://dx.doi.org/10.1038/nprot.2007.132.

35. Yang F, Zhang S, Zhou YJ, Zhu Z, Lin X, Zhao ZK. 2012. Character-ization of the mitochondrial NAD�-dependent isocitrate dehydrogenaseof the oleaginous yeast Rhodosporidium toruloides. Appl Microbiol Bio-technol 94:1095–1105. http://dx.doi.org/10.1007/s00253-011-3820-3.

36. Huh W-K, Falvo JV, Gerke LC, Carroll AS, Howson RW, Weissman JS,O’Shea EK. 2003. Global analysis of protein localization in budding yeast.Nature 425:686 – 691. http://dx.doi.org/10.1038/nature02026.

37. Jiang H, He J, Pu S, Tang C, Xu G. 2007. Heat shock protein 70 istranslocated to lipid droplets in rat adipocytes upon heat stimulation.Biochim Biophys Acta 1771:66 –74. http://dx.doi.org/10.1016/j.bbalip.2006.10.004.

38. Zhang H, Wang Y, Li J, Yu J, Pu J, Li L, Zhang H, Zhang S, Peng G,Yang F, Liu P. 2011. Proteome of skeletal muscle lipid droplet revealsassociation with mitochondria and apolipoprotein A-I. J Proteome Res10:4757– 4768. http://dx.doi.org/10.1021/pr200553c.

39. Li Z, Thiel K, Thul PJ, Beller M, Kuhnlein RP, Welte MA. 2012. Lipiddroplets control the maternal histone supply of Drosophila embryos. CurrBiol 22:2104 –2113. http://dx.doi.org/10.1016/j.cub.2012.09.018.

40. Wang C, St Leger RJ. 2007. The Metarhizium anisopliae perilipin ho-molog MPL1 regulates lipid metabolism, appressorial turgor pressure,and virulence. J Biol Chem 282:21110 –21115. http://dx.doi.org/10.1074/jbc.M609592200.

41. Naested H, Frandsen GI, Jauh GY, Hernandez-Pinzon I, Nielsen HB,Murphy DJ, Rogers JC, Mundy J. 2000. Caleosins: Ca2�-binding pro-teins associated with lipid bodies. Plant Mol Biol 44:463– 476. http://dx.doi.org/10.1023/A:1026564411918.

42. Jiang PL, Chen JC, Chiu ST, Tzen JT. 2009. Stable oil bodies sheltered bya unique caleosin in cycad megagametophytes. Plant Physiol Biochem47:1009 –1016. http://dx.doi.org/10.1016/j.plaphy.2009.07.004.

43. Poxleitner M, Rogers SW, Lacey Samuels A, Browse J, Rogers JC. 2006.A role for caleosin in degradation of oil-body storage lipid during seedgermination. Plant J 47:917–933. http://dx.doi.org/10.1111/j.1365-313X.2006.02845.x.

44. Brodhun F, Gobel C, Hornung E, Feussner I. 2009. Identification ofPpoA from Aspergillus nidulans as a fusion protein of a fatty acid hemedioxygenase/peroxidase and a cytochrome P450. J Biol Chem 284:11792–11805. http://dx.doi.org/10.1074/jbc.M809152200.

45. Tsitsigiannis DI, Zarnowski R, Keller NP. 2004. The lipid bodyprotein, PpoA, coordinates sexual and asexual sporulation in Aspergil-lus nidulans. J Biol Chem 279:11344 –11353. http://dx.doi.org/10.1074/jbc.M310840200.

46. Boer VM, Crutchfield CA, Bradley PH, Botstein D, Rabinowitz JD.2010. Growth-limiting intracellular metabolites in yeast growing underdiverse nutrient limitations. Mol Biol Cell 21:198 –211. http://dx.doi.org/10.1091/mbc.E09-07-0597.

47. Athenstaedt K, Zweytick D, Jandrositz A, Kohlwein SD, Daum G. 1999.Identification and characterization of major lipid particle proteins of theyeast Saccharomyces cerevisiae. J Bacteriol 181:6441– 6448.

48. Aime MC, Matheny PB, Henk DA, Frieders EM, Nilsson RH, Piepen-bring M, McLaughlin DJ, Szabo LJ, Begerow D, Sampaio JP, Bauer R,Weiss M, Oberwinkler F, Hibbett D. 2006. An overview of the higherlevel classification of Pucciniomycotina based on combined analyses ofnuclear large and small subunit rDNA sequences. Mycologia 98:896 –905.http://dx.doi.org/10.3852/mycologia.98.6.896.

49. Nguyen LN, Hamari Z, Kadereit B, Trofa D, Agovino M, Martinez LR,Gacser A, Silver DL, Nosanchuk JD. 2011. Candida parapsilosis fat stor-age-inducing transmembrane (FIT) protein 2 regulates lipid droplet for-mation and impacts virulence. Microbes Infect 13:663– 672. http://dx.doi.org/10.1016/j.micinf.2011.02.009.

50. Buzzini P, Innocenti M, Turchetti B, Libkind D, van Broock M, Muli-nacci N. 2007. Carotenoid profiles of yeasts belonging to the genera Rho-dotorula, Rhodosporidium, Sporobolomyces, and Sporidiobolus. Can J Mi-crobiol 53:1024 –1031. http://dx.doi.org/10.1139/W07-068.

51. Ageitos JM, Vallejo JA, Veiga-Crespo P, Villa TG. 2011. Oily yeasts as

Dynamics of R. toruloides Lipid Droplet Proteome

March 2015 Volume 14 Number 3 ec.asm.org 263Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

oleaginous cell factories. Appl Microbiol Biotechnol 90:1219 –1227. http://dx.doi.org/10.1007/s00253-011-3200-z.

52. Yang H, Galea A, Sytnyk V, Crossley M. 2012. Controlling the size oflipid droplets: lipid and protein factors. Curr Opin Cell Biol 24:509 –516.http://dx.doi.org/10.1016/j.ceb.2012.05.012.

53. Fei W, Shui G, Zhang Y, Krahmer N, Ferguson C, Kapterian TS, LinRC, Dawes IW, Brown AJ, Li P, Huang X, Parton RG, Wenk MR,Walther TC, Yang H. 2011. A role for phosphatidic acid in the formationof “supersized” lipid droplets. PLoS Genet 7:e1002201. http://dx.doi.org/10.1371/journal.pgen.1002201.

54. Fei W, Li H, Shui G, Kapterian TS, Bielby C, Du X, Brown AJ, Li P,Wenk MR, Liu P, Yang H. 2011. Molecular characterization of seipin andits mutants: implications for seipin in triacylglycerol synthesis. J Lipid Res52:2136 –2147. http://dx.doi.org/10.1194/jlr.M017566.

55. Kadereit B, Kumar P, Wang WJ, Miranda D, Snapp EL, Severina N,Torregroza I, Evans T, Silver DL. 2008. Evolutionarily conserved genefamily important for fat storage. Proc Natl Acad Sci U S A 105:94 –99.http://dx.doi.org/10.1073/pnas.0708579105.

56. Froissard M, D’Andrea S, Boulard C, Chardot T. 2009. Heterologousexpression of AtClo1, a plant oil body protein, induces lipid accumulationin yeast. FEMS Yeast Res 9:428 – 438. http://dx.doi.org/10.1111/j.1567-1364.2009.00483.x.

57. Shi S, Valle-Rodriguez JO, Khoomrung S, Siewers V, Nielsen J. 2012. Func-tional expression and characterization of five wax ester synthases in Saccharomy-ces cerevisiae and their utility for biodiesel production. Biotechnol Biofuels 5:7.http://dx.doi.org/10.1186/PREACCEPT-1932279820621895.

58. Jin G, Zhang Y, Shen H, Yang X, Xie H, Zhao ZK. 2013. Fatty acid ethylesters production in aqueous phase by the oleaginous yeast Rhodospo-ridium toruloides. Bioresour Technol 150:266 –270. http://dx.doi.org/10.1016/j.biortech.2013.10.023.

59. Bajaj A, Lohan P, Jha PN, Mehrotra R. 2010. Biodiesel productionthrough lipase catalyzed transesterification: an overview. J Mol Catal BEnzym 62:9 –14. http://dx.doi.org/10.1016/j.molcatb.2009.09.018.

60. Beller M, Riedel D, Jansch L, Dieterich G, Wehland J, Jackle H,Kuhnlein RP. 2006. Characterization of the Drosophila lipid droplet sub-

proteome. Mol Cell Proteomics 5:1082–1094. http://dx.doi.org/10.1074/mcp.M600011-MCP200.

61. Kuerschner L, Moessinger C, Thiele C. 2008. Imaging of lipid biosyn-thesis: how a neutral lipid enters lipid droplets. Traffic 9:338 –352. http://dx.doi.org/10.1111/j.1600-0854.2007.00689.x.

62. Gangar A, Karande AA, Rajasekharan R. 2001. Isolation and local-ization of a cytosolic 10 S triacylglycerol biosynthetic multienzymecomplex from oleaginous yeast. J Biol Chem 276:10290 –10298. http://dx.doi.org/10.1074/jbc.M009550200.

63. Guo Y, Walther TC, Rao M, Stuurman N, Goshima G, Terayama K,Wong JS, Vale RD, Walter P, Farese RV. 2008. Functional genomicscreen reveals genes involved in lipid-droplet formation and utilization.Nature 453:657– 661. http://dx.doi.org/10.1038/nature06928.

64. Beller M, Sztalryd C, Southall N, Bell M, Jackle H, Auld DS, Oliver B.2008. COPI complex is a regulator of lipid homeostasis. PLoS Biol 6:e292.http://dx.doi.org/10.1371/journal.pbio.0060292.

65. Soni KG, Mardones GA, Sougrat R, Smirnova E, Jackson CL, Bonifa-cino JS. 2009. Coatomer-dependent protein delivery to lipid droplets. JCell Sci 122:1834 –1841. http://dx.doi.org/10.1242/jcs.045849.

66. Bostrom P, Andersson L, Rutberg M, Perman J, Lidberg U, JohanssonBR, Fernandez-Rodriguez J, Ericson J, Nilsson T, Boren J, Olofsson SO.2007. SNARE proteins mediate fusion between cytosolic lipid droplets andare implicated in insulin sensitivity. Nat Cell Biol 9:1286 –1293. http://dx.doi.org/10.1038/ncb1648.

67. Hutagalung AH, Novick PJ. 2011. Role of Rab GTPases in membranetraffic and cell physiology. Physiol Rev 91:119 –149. http://dx.doi.org/10.1152/physrev.00059.2009.

68. Martin S, Driessen K, Nixon SJ, Zerial M, Parton RG. 2005. Regulatedlocalization of Rab18 to lipid droplets: effects of lipolytic stimulation andinhibition of lipid droplet catabolism. J Biol Chem 280:42325– 42335.http://dx.doi.org/10.1074/jbc.M506651200.

69. Rodriguez-Navarro JA, Cuervo AM. 2010. Autophagy and lipids: tight-ening the knot. Semin Immunopathol 32:343–353. http://dx.doi.org/10.1007/s00281-010-0219-7.

Zhu et al.

264 ec.asm.org March 2015 Volume 14 Number 3Eukaryotic Cell

on Septem

ber 24, 2020 by guesthttp://ec.asm

.org/D

ownloaded from